Anal. Chem. 1991, 63, 2901-2965
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Enzyme-Modified Organic Conducting Salt Microelectrode Jodi L. Kawagoe, David E. Niehaus, and R. Mark Wightman* Department of Chemistry, University of North Carolina at Chapel Hill, Chapel Hill, North Carolina 27599-3290 A miniaturized enzyme-modified electrode has been constructed and evaluated. The tip of a capiiiaryancased, carbon-fiber electrode is recessed, and tetrathlafuivaiene-tetracyanoqulnodlmethanecrystals are electrochemically deposited In the recessed tip. Fiavoenzymes are placed in the recess by cross-linking with glutaraldehyde, The specHic enzymes used are glucose oxidase to form a microbiosensor for glucose, and a combination of acetylcholine esterase and choline oxidase to form a microbiosensor for acetylcholine. The sensor is operated in an amperometric mode with E, = 150 mV versus a sodium saturated calomel electrode, and the response appears to be limited by the kinetics of the enzyme reaction. The effective maximum current density for the glucose electrode is greater than 600 pA/cm*. At low concentrations of glucose, oxygen provides a significant interference by attenuating the signal. The device is simple to prepare and has a rapid response time. Interference from ascorbate has been significantly reduced by the design and by addition of a layer of ascorbate oxidase. Aithough not yet suitable for use in tissue, the biosensors are suitable for detection in sltuatlons where oxygen concentrations do not frequently change.
INTRODUCTION Amperometric biosensors based on redox enzymes have been the subject of considerable recent interest, and their principles have been the subject of recent reviews ( I , 2). In general these devices achieve specificity for the substrate of interest via the enzyme which is immobilized on the electrode surface. Current which results from the enzyme-mediated reaction is measured, and is proportional to the substrate concentration. Because electron transfer between metal or carbon electrodes and enzymes is generally quite slow, several schemes have been developed to catalyze this process. Biosensors for the detection of glucose illustrate many of the approaches that have been taken in the construction of amperometric biosensors. Glucose oxidase (GOX), a flavoenzyme, is immobilized at an electrode surface by a membrane or with a cross-linking agent such as glutaraldehyde. This enzyme is specific for the ,f3 anomer of D-glucose and catalyzes the overall ricction P-D-glucose
+ O2 9 H20
GOX
D-gluconic acid 9 H202
Traditionaily, the enzyme reaction has been monitored by detecting the formation of peroxide or the consumption of oxygen ( 3 , 4 ) . Chemical interference can arise from species which are electroactive at the applied potential (3) or from limited oxygen availability which can iower the reaction rate arid, thus, the measured current. To overcome the problem of oxygen availability, soluble eiectron-transfer media" have been developed which couple eiectrori transfer between the electrode and enzyme (5). These reagents are retained a t the electrode surface with a membrane, or are entrapped in an eiectrogenerated film a t the surface (6, 7). Alternativeiy, the mediator can be kept in the
* To whom correspondence
should be addressed. 0003-2700/91/0363-2961$02.50/0
interfacial region by attachment to a polymer chain (8) or incorporation into the immobilized enzyme (9). Another approach is based on the observation that electrode surfaces formed from conducting organic salts, such as tetrathiafulvalene-tetracyanoquinodimethane (TTF-TCNQ), permit electron transfer with a variety of redox enzymes without added mediator (10-14). The mechanism of electron transfer is not understood (15, 16) but may involve homogeneous mediation by dissolved electrode components (16, 17),heterogeneous redox catalysis involving a mobile surface species, or direct electron transfer (18). An advantage of organic salt electrodes is that enzymes can be adsorbed by simply soaking in an aqueous solution of the enzyme (19,201. With all of these schemes the sensor can function well in the absence of oxygen, but high oxygen concentrations can provide interference by competing with the mediated reaction. An amperometric biosensor suitable for measurements in tissue requires high chemical specificity and must be of small size to provide minimal damage (21,22). In this paper we describe an approach to the construction of a miniature biosensor with an outside diameter of approximately 20 pm based on a carbon-fiber electrode (23). The carbon tip has been recessed to allow incorporation of the biosensor elements. Recessed microelectrodes offer the advantage that the biosensor components are protected in the recess (24). TTFTCNQ is placed in the recess by electrocrystallization, and the recess is filled with a gel matrix of the enzyme of interest made by cross-linking with glutaraladehyde (25). The versatility of the electrode design is demonstrated by the detection of glucose, when the electrode contains glucose oxidase, and acetylcholine with the use of acetylcholine esterase and choline oxidase. In addition, the effects of various chemical interferences are evaluated.
EXPERIMENTAL SECTION Reagents. Tetrathiafulvalene (TTF) and lithium iodide were obtained from Aldrich (Milwaukee,WI). Acetonitrile (UV grade) was from Burdick and Jackson (Muskegon, MI). All other chemicals were from Sigma Chemical Co. (St. Louis, MO) and were used as received. The enzymes employed were glucose oxidase (EC 1.1.3.4; type X-Sfrom Aspergillus niger; activity 1500oO units/g), ascorbate oxidase (EC 1.10.3.3; from Cucurbita species; activity 1700 unita/mg), acetylcholine esterase (EC 3.1.1.7; type I11 from electric eel, aqueous solution containing approximately 5 mg of (NH4)$04/mg of protein; activity 10oO units/mg protein), and choline oxidase (EC 1.1.3.17; activity 10.7 units/mg, from Alcaligenes species). Tetrathiafulvalene-tetracyanoquinodimethane (TTF-TCNQ) crystals (26) and Li-TCNQ (27) were prepared according to literature procedures. Doubly distilled water was used in all aqueous solutions. Unlesa noted, all solutions were prepared in 0.1 M Na2HP0,, 0.1 M NaCl adjusted to pH 7.4 with HCl. Glucose stock solutions (1M) were allowed to mutorotate for at least 24 h before use to establish equilibrium concentrations of a and @ anomers and were stored at 4 "C. Reported concentrations are those of total glucose although glucose oxidase is specific for @-glucose.Substrate concentrations were made from the same stock solution. Electrode Preparation. Carbon-fiber microelectrodes were made with Thornel T300 carbon fibers (7-pm diameter, Amoco Performance Products, Inc., Greenville, SC) sealed into tapered glass capillaries (23) with epoxy (Epon 828 with 14% metaphenylenediamine, Miller Stephenson Chemical Co., Danbury, CT). The epoxy was cured at 180 "C for at least 1h after sitting at room temperature overnight. The electrodes were bevelled at 0 1991 American Chemical Society
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ANALYTICAL CHEMISTRY, VOL. 63, NO. 24, DECEMBER 15, 1991
a 45" angle on a micropipet beveler (Model BV-10, Sutter Instrument Co., Novato, CA). The carbon fiber was recessed into the glass capillary by electrochemically etching the tip of the electrode in a solution containing 0.5 mM potassium dichromate, 5 M sulfuric acid, and a Pt counter electrode. A 20-V peak-topeak, 60-Hz sine wave applied with a Krohn-Hite 1200A function generator (Avon, MS) for 1 min yielded recessed carbon cones whose tips were 10-40 pm from the end of the bevelled glass. ?TF-TCNQ crystals were galvanostatically deposited (0.13 pA) in the recessed tip from an acetonitrilesolution containing 30 mM TTF and saturated with both TTF-TCNQ and Li-TCNQ (28). A platinum wire was used as the cathode. After approximately 2 min TTF-TCNQ crystals extended from the recessed tip. Crystals external to the capillary were gently removed with a soft tissue. Glucose oxidase was added to the electrode tip by cross-linking glucose oxidase with glutaraldehyde. The TTF-TCNQ microelectrodes were soaked for 1-3 min in 80 pL of aqueous buffer containing 5 mg of glucose oxidase (750 units) and 3 pL of 25% aqueous glutaraldehyde (29). Electrodes were dried for 3 h in a desiccator and were stored in buffer (pH 7.4) at 4 "C when not in use. In some cases a layer of cross-linked ascorbate oxidase (AAO) was placed over the cross-linked glucose oxidase layer. AAO was cross-linked by adding 3 pL of 25% aqueous glutaraldehyde to a vial containing lo00 units of AAO in 80 pL of buffer and mixing thoroughly. The TTF-TCNQ microelectrodes containing the cross-linked glucose oxidase were soaked in this solution for a few minutes and dried in a desiccator for 3 h. Electrodes suitable for detection of acetylcholine (20) were prepared by immobilizing acetylcholine esterase and choline oxidase in the recess of the TTF-TCNQ microelectrodes. A mixture of 125 p L of acetylcholine esterase (125 units) and choline oxidase (125units) was shaken well and sonicated. An 8pL aliquot of 25% aqueous glutaraldehyde was added and mixed thoroughly. The electrodes were soaked in the enzyme solution for 1-3 min and dried in a desiccator for 3 h. Electrodes were stored in buffer (pH 7.4) at 4 "C. Instrumentation. Light micrographs of electrodes were taken using a Zeiss Axiovert 35 microscope (Carl Zeiss, Inc., Southern Microscope, Raleigh, NC). The potentiostat for electrochemical studies was a BAS CV-37 (Bioanalytical Systems, West Lafayette, IN) used with a Model 2000 XY recorder or an Omniscribe strip chart recorder (both from Houston Instrument, Austin, TX). The reference electrode was a saturated sodium calomel electrode (SSCE) with a platinum-wire counter electrode. Enzyme-modified electrodes were evaluated by insertion of the tip into the outlet of a flow injection analysis system constructed with glass-lined stainless steel tubing. In all experiments, the buffer was deoxygenated. Analytes were introduced with a loop injector (Rheodyne Model 7010, Cotati, CA) with a 500-pL sample loop. The pump (ISCO pLC-500 micropump, Lincoln,NE)was operated at 250 pL/min. Experiments were performed at 23 "C. Kinetic Analysis. Current versus concentration data were analyzed in order to see if the overall electrode kinetics followed an electrochemical form of the Michaelis-Menten equation: . . C z = I,,(1) C + KM' The apparent Michaelis constant,KM', of the immobilized enzymes was determined with a form of the EadieHoftsee equation (30): . . - &'(i/C) z ,=, z (2) where i is the current,,,i is the maximum current when the enzyme is saturated, and C is the substrate concentration. Equation 1 is appropriate when the enzyme kinetics are rate limiting. If diffusion is rate limiting, the KM' may be considerably larger (31)than the intrinsic K M of the enzyme, and the plots of eq 2 will be nonlinear when a sufficientlylarge concentration range is explored.
RESULTS AND DISCUSSION Characterization of Recessed Electrodes. Light micrographs taken at each stage in the fabrication of a recessed TTF-TCNQ microelectrode are shown in Figure 1. Parts A and B of Figure 1show a carbon-fiber microelectrode before
B
c
D
Figure 1. Light micrographs illustrating steps in the fabrication of a recessed lT-TCNQ microelectrode: (A) carbon-fiber microelectrode; (B) etched carbon-fiber microelectrode; (C) electrode after electrocrystallization of lTF-TCNQ in etched carbon-fiber microelectrode; (D) recessed TTF-TCNQ microelectrode. Note: carbon-fiber diameter is 7 pm. 0.4
1 :::
i (nA)
I
-
0
.
4
1
400
"
'
200
"
'
0
E (mV v s SSCE)
'
'
.
I
200
'
-0.1
0
E (mV va SSCE)
Figure 2. Cyclic voltammograms recorded at a etched carbon-fiber microelectrode(A) and the same electrode following electrocrystallization of TTF-TCNQ (B). Scan rate: 50 mV/s. Solution conditions: (A) 1 mM Fe(CN),%, 0.1 M K2S04,pH 3.0; (B) 1 mM Fe(CN),%, pH 7.4.
and after the etching procedure. The electrode after TTFTCNQ deposition is shown in Figure 1C; TTF-TCNQ crystals can be seen extending from the recess opening. Excess crystals have been removed in Figure 1D so that all remaining crystals are in the recess. Cyclic voltammograms recorded in Fe(CN),& solutions (pH 3.0) with recessed electrodes (Figure 1B) show large background currents and peaks for the redox processes (Figure 2A), the behavior expected at recessed microelectrodes for faradaic processes dominated by linear diffusion. When the recess is partially filled with TTF-TCNQ crystals (Figure lD), a sigmoidal voltammograms are obtained in ferricyanide solutions (pH 7.4) with much lower charging current (Figure 2B), indicating that the recess is a t least partially filled. At carbon electrodes the electrochemical rate of the reduction of ferricyanide at pH 7.4 is very slow (32),resulting in drawn out voltammograms. The well-defined voltammograms obtained in this solution after deposition indicate that the faradaic current is largely due to electron transfer a t the TTF-TCNQ crystals. Although the microscopically observed area of the TTF-TCNQ crystals appears large, the steadystate limiting current is much smaller than at the carbon-fiber microelectrode before etching. This behavior is expected for electrodes with a relatively shallow recess (33).
Recessed Electrodes Coated with Glucose Oxidase. The amperometric responses to deaerated solutions of 1and 10 mM glucose for TTF-TCNQ microelectrodes containing cross-linked glucose oxidase are shown in Figure 3. Noise levels are less than 0.2 PA. The response time to a bolus of glucose, defined as the time for the current to rise to 90% of its maximum, was less than 1.5 s. Typically, the response would decrease by approximately 20% in the first hour of operation and then remain stable with continued operation over the course of several hours.
ANALYTICAL CHEMISTRY, VOL. 63,NO. 24, DECEMBER 15, 1991
M
H 5 min
1 mM glucose
0.0
10 mM glucose
Table I. Average Currents Recorded at TTF-TCNQ Electrodes Coated with Glucose Oxidase at E,, = 150 m V (Errors Given as Standard Deviations)
packed 420 f 240 TTF-TCNQ crystals, r = 120 @ma TTF-TCNQ microelectrodea TTF-TCNQ microelectrodeb
17 f 13 178 f 86
1.0
* 06
h
A
4
4
A
100 200 E (mV vs SSCE)
0
,
10 mM glucose i, pA j , pA/cm2
Air
b
Flgurr 3. Current response to glucose in a flow inJection apparatus of a m-TCNQ microelectrodecoated with glutaraldehydecross-Unked glucose oxidase. Results were obtained with 500-pL injections of 1 and 10 mM glucose in nitrogen-saturated soiutlons. E = 4-150 mV vs SCE flow rate = 250 pL/min.
electrode type
A
N2
O
2963
300
Flgurr 4. Potential dependence of glucose (1 mM) oxidation at TTFTCNQ microelectrodes coated with cross-linked glucose oxidase de-
termined in a flow injection apparatus: (circles) nitrogen-saturated soiutbn; (triangles) air-saturated soiution. The current was normalized to the nitrogen-saturated response at +300 mV. Data represent the average of five electrodes, with error bars representing one standard deviation.
0.3 mM ascorbic acid i , pA j , fiA/cma i
13,000f 800 31 f 19 i m .
31 f 24
35 f 9
65 f 16
330 f 160 25 f 9
46 f 16
aGlucose oxidase (1 mg/mL) adsorbed onto the electrode by overnight soaking. *Glucose oxidase cross-linked with glutaraldehyde. The effective current density a t TTF-TCNQ electrodes coated with glutaraldehyde cross-linked glucose oxidase is much greater than electrodes at which the enzyme is simply adsorbed (Table I). This was true when compared to electrodes formed from packed crystals or to electrodeposited TTF-TCNQ microelectrodes. The response of the electrodes with adsorbed glucose oxidase also deteriorated more rapidly than the "F-TCNQ microelectrodes coated with cross-linked glucose oxidase. This has been attributed to loss of active enzyme from the interfacial region (25). The potential dependence of the amperometric response was tested with 1 mM glucose in deaerated solutions over the range 0.0-300 mV. The current increased linearly with potential (Figure 41,consistent with the results of Hale and Skotheim (16) obtained with adsorbed glucose oxidase on TTF-TCNQ electrodes. The amperometric response to 5 mM glucose changed by less than 1 % with flow rates from 0.250 to 1.00 mL/min.
Kinetics of TTF-TCNQ Microelectrodes Coated with Glucoae Oxidase. The kinetic behavior of these electrodes was evaluated from the amperometric response (Eapp= 150 mV) to concentrations of glucose varying from 1to 40 mM. Plots of the data using eq 2 were linear, and the apparent KM' value was 2.5 f 0.9 mM (n = 4 electrodes). Calibration curves and the curve from eq 1 using this KM' model are shown in Figure 5A. The intrinsic K M for the glucose oxidase preparation used in this work was not determined; however, Kamin and Wilson (34) reported that type I1 glucose oxidase from Sigma has a K M of 6.8 mM. The linear plots of eq 2 combined with KM' < K M and the flow-rate independence all indicate that the response is not limited by diffusion. The average, i was 0.42 f 0.09 nA at E,, = 150 mV, which, on the basis of the area of the hole at the tip of the recess, gives an estimated
0.00 1 .oo
i ( a i r ) 0.75 i (N,)
0.50 0.25 0.00 0
10
20
30
40
50
[Glucosa] (mM)
current as a function of &cose conmtration measured at TTF-TCNQ microelectrodes coated with giutaraidehyde cross-linked glucose oxidase: (circles)results from N,-saturated solutions; (triangles) results from air-saturated solutions; (solid line) MIchaelis-Menten equation using K,,,' and, / determined from EadieHoftsee plot (correlation coefficient with data is 0.993). The results are averages from fwelectrodes, and each response was "allzed to the respective i,. E, = +150 mV. (B) Ratio of currents in air to that measured in N,-saturated solutions as a function of glucose concentration for three amperomeMc blosensors: (squares)this work; (triangles)from Hale et ai. (35);(circles)from &egg and Heiler (36). Figwe 6. (A) "allzed
limiting current density,,j in the range 600-1000 pA/cm2. This indicator of sensitivity compares favorably with other enzyme-based electrodes. Hale et al. (35)and Gregg and Heller (36)reported glucose electrodes with j-'s of 29-275 and 100-800 pA/cm2 based on a polymer-tethered mediator and a conducting polymer, respectively. High current densities are especially desirable for a miniature sensor such as described here to ensure a detectable signal. The apparent KM' with E, = 300 mV was 14 mM, with an 8-fold increase in, i over that found at E,, = 150 mV. Thus, it appears that the electrode is diffusionally controlled a t the higher applied potential. Diffusionally limited conditions are advantageous because the linear range of the enzyme electrode is increased as a result of the larger K M However, to minimize interference from easily oxidized substances such as ascorbate, all of the remaining experiments were conducted with Eapp= 150 mV. Interference by Oxygen. Oxygen is the natural oxidant for glucose oxidase, which has a KM of 480 pM for this substrate (37), and it can compete with the electrode as the electron sink and attenuate the amperometric signal. Indeed, oxygen must be evaluated as a potential interference at any glucose oxidase based sensor that is to be used in environments
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ANALYTICAL CHEMISTRY, VOL. 63,NO. 24, DECEMBER 15, 1991
where changes in oxygen concentration can occur. Oxygen concentrations can very rapidly and by a considerable amount in biological tissue (38). An attenuation of the amperometric current for 1 mM glucose was seen at all potentials, but the interference was less a t the more positive potential (Figure 4). Presumably, this reflects the increased ability of the electrode to compete with oxygen as the sink for electrons. Interference by oxygen is less at higher glucose concentrations (Figure 5B). In this figure results from this work are compared to results reported by Hale et al. (35)and Gregg and Heller (36). While all three types of electrodes are sensitive to oxygen, the electrodes described here are superior in the concentration range of physiological interest, between 1and 10 mM (39). Note that interference has been measured for the largest possible change in oxygen concentration under physiological conditions, from negligible oxygen to air-saturated conditions (approximately 240 p M oxygen). The range of oxygen levels in the rat brain measured with implanted electrodes is C5-50 pM (38),and thus interference in vivo will be less than seen in this figure. Interference by Ascorbate. Another possible interference is ascorbic acid which is electroactive in the applied potential range, and which is present at concentrations of greater than 200 pM in the extracellular fluid of the brain (40,41). The amperometric responses of different TTF-TCNQ electrodes coated with glucose oxidase to 300 pM ascorbate in nitrogen-saturated solutions are also given in Table I. The current density for ascorbate oxidation is similar a t all three types of electrodes and is less than measured a t a bevelled carbon-fiber electrode. This suggests that the amperometric response a t this potential is under electrode-kinetic control. The responses summarized in Table I show that the interference from ascorbate is less of a problem when efficient catalysis of glucose oxidation is achieved. In an effort to further attenuate the ascorbate signal, a layer of glutaraldehyde cross-linked ascorbate oxidase was placed over the cross-linked glucose oxidase layer. AAO catalyzes the reaction
AAO
2 L-ascorbate + O2 2 dehydroascorbate + 2 H 2 0 By placing this layer over the glucose oxidase layer, TTFTCNQ should not interact with the ascorbate oxidase. This treatment did not affect the amplitude of the amperometric response to ascorbate in deaerated solutions. However, in air-saturated solutions the response to ascorbate was completely eliminated. Thus, the ascorbate oxidase catalyzes the mutual destruction of two major interferents. At these double-coated electrodes the response to 1 mM glucose in airsaturated solutions was larger in the presence of 300 pM ascorbate by a factor of 1.9 f 0.3 (average of four electrodes). While this response still indicates an interference by oxygen, it does provide some improvement. Uric acid was also tested as a possible interferent. No response was observed to injections of 190 pM uric acid. Acetylcholine Microelectrode. Electrodes responsive to acetylcholine were prepared with the TTF-TCNQ microelectrodes coated with choline oxidase and acetylcholine esterase (20). Acetylcholine esterase (AChE) catalyzes the reaction
AChE
acetylcholine + H 2 0 acetate + choline Choline oxidase (ChO), a flavoenzyme, converts choline to betaine in two steps choline
+ O2
ChO
+ H20z betaine + Hz02
betaine aldehyde
ChO
betaine aldehyde + O2 At TTF-TCNQ electrodes the requirement for oxygen is short-circuited. The amperometric responses for 5 and 800
T
0 . 2 PA
5 pM Acetylcholine
~
8 0 0 1 M Acetylcholine
Figure 6 . Current response to acetylcholine of TTF-TCNQ microelectrodes coated with cross-linked choline oxidase and acetylcholine esterase. The results are for 5 and 800 pM acetylcholine injections in nitrogen-saturated solutions. Other conditions are as in Figure 3.
pM acetylcholine are shown in Figure 6.
The RMS noise ranged from 15 to 50 fA. Unlike the glucose electrode, oxygen provides less of an interference a t low concentrations. The response time to acetylcholine is less than 4 s. Values of KM' were determined using eq 2 with data obtained in nitrogen-saturated solutions and had correlation coefficients of approximately 0.9. The apparent KM' found for acetylcholine was 150 f 50 pM (n = 4) which is close to that found with AChE and ChO adsorbed on TTF-TCNQ crystals (20). This value indicates a kinetically controlled response, and consistent with this, identical KM' values were obtained in quiescent solutions as in the flow injection apparatus. The response to ascorbate with these electrodes was identical to that found with the glucose oxidase electrodes. Since the response to acetylcholine is much smaller, ascorbate did cause significant interference. Registry No. Glucose, 50-99-7; acetylcholine, 51-84-3;glycose oxidase, 9001-37-0; acetylcholine esterase, 9000-81-1; choline oxidase, 9028-67-5; tetrathiafulvalene, 31366-25-3; tetracyanoquinodimethane, 1518-16-7; ascorbic acid, 50-81-7; oxygen, 7782-44-7; ascorbate oxidase, 9029-44-1. LITERATURE CITED Frew, J. E.; Hill, H. A. 0. Anal. Chem. 1987, 59, 933A-944A. Heller, A. Acc. Chem. Res. 1990, 23, 128-134. Harrison, D. J.; Turner, R . F. B.; Bakes, H. P. Anal. Chem. 1988, 6 0 , 2002-2007. Shimuzu, Y.; Morita, K. Anal. Chem. 1990, 6 2 , 1498-1501. Aston, W. J.; Cass, A. E. G.;Davis, G.;Francis, G. D.; Hill, H. A. 0.; Hiaains. I. J.; Plotkln, E. V.; Scott, L. D. L.; Turner, A. P. F. Anal. C&m. 1984, 56, 667-671. Umana. M.; Waller, J. Anal. Chem. 1986, 5 8 , 2979-2983. Malitesta, C.; Palmisano, F.; Torsi, L.; Zambonin, P. G. Anal. Chem. 1QQO.62. ~. 2735-2740. Hale, P. D.; Inagaki, T.; Karan, H. I . ; Okamoto, Y.; Skotheim, T. A. J . Am. Chem. Soc. 1989, 1 1 1 , 3482-3484. Degani, Y.; Heller. A. J. Phys. Chem. 1987, 91, 1285-1289. Kulys, J. J.; Samalius, A. S.; Svirmickas, G.J. S. FEBS Lett. 1980, f. l.d. , 7-10 . .- . Albery, W. J.; Bartlett, P. N.; Craston, D. H. J. Electroanal. Chem. lQn5. 194. 223-235. Ai&&, W.' J : & k t , P. N.; Bycrofl, M.; Craston, D. H.; Drlscoll, B. J. J. Nectroanal. Chem. 1987, 218, 119-126. McKenna, K.; Brajter-loth, A. Anal. Chem. 1987, 59, 954-958. Zhao, S.;Lennox, R. 8. Anal. Chem. 1991, 6 3 , 1174-1178. Hill, B. S.; Scolari, C. A.; Wilson, G. S.J . Necfroanal. Chem. Interfacial Necfrochem. 1988, 252, 126-138. Hale, P. D.; Skotheim, T. A. Synth. Met. 1989, 2 8 , C853-C858. Cenas. N. K.; Kulys, J. J. Bioelectrochem. Bioenerg. 1981, 8 , 103-1 13. Albery, W. J.; Bartlett, P. N.; Cass, A. E. G. Phil. Trans. R. SOC. London B 1987, 316, 107-119. Boutelle, M. G.;Standford, C.; Fillenz, M.; Aibery, W. J.; Bartlett, P. N. Neurosci. Lett. 1988, 72, 283-288. Hale, P. D.; Wightman, R. M. Mol. Cysf. L i q . Crysf. 1988, 160, 269-279. Ikariyama, Y.; Yamauchi, S.; Yukiashi, T.; Ushioda, H. J. Electfochem. SOC. 1989, 136, 702-706. Kuhr, W. G.; Pantano. P.; Morton, T. H. J. Am. Chem. SOC. 1991, 113. 1832-1833. Kelly, R.; Wightman, R. M. Anal. Chem. Acta 1988, 187, 79-87. Morita. K.; Shimuzu, Y. Anal. Chem. 1989, 6 1 , 159-162.
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RECEIVED for review June 17,1991. Accepted September 25, 1991. This research was supported by NIH (Grant NS15841).
Strategies for Low Detection Limit Measurements with Cyclic Voltammetry Donna J. Wiedemann, Kirk T. Kawagw, Robert T. Kennedy, Edward L. Ciolkowski, and R. Mark Wightman* Department of Chemistry, University of North Carolina, Chapel Hill, North Carolina 27599-3290
Cyclk voltammetry of Naflon-coated, carbon-flber electrodes lo used to detect trace concentrations of dopamine, both In a flow Injectbn apparatus and In the kaln of an anaesthetlzed rat. To Improve slgnal-to-noise ratios, the sources of nolse durlng cyclk voltammetry have been determined and strategies have been developed to decrease the noise. With the potentlostat employed, the measured noise is comparable to that expected for Johnson noise from the feedback reslstor of the current transducer. Addltional nolse arlses from the wavefon generator empbyed and, in same cases, lIne nolse. Line nolse Is dlscrimlnated against by starting each cyclic voltammogram elther In phase or 180' out of phase wlth the ihre frequency. When used In vlvo, addltknal ndse also arises from the physlologlcal actlvlty of the anlmal. Detection limlts are found to closely correspond to those predlcted on the basis of slmulatlon of the voltammetric shape and the measured nolse. Detection limits are improved by the use of appropriate analog and dlgltal fllterlng, ensemble averaging, and appropriate timlng of repetltlve cycllc voltammograms. The combined use of these technlques enables the In vlvo detectlon of approximately 100 nM of dopamine with a slgnal-to-noise ratio of 25.
INTRODUCTION The increased use of voltammetry for in vivo detection of trace concentrations of catecholamine neurotransmitters has led us to explore techniques to improve signal-to-noise ratios in these experiments. Signal-processing techniques such as ensemble averaging (1-4) and the use of analog and digital filters (5-8) have long been employed to improve the quality of a variety of types of data in analytical chemistry. However, these techniques have been less frequently used with voltammetric methods to enhance signal detection. Smith used on-line Fourier transformation to digitally smooth voltammetric data (9, 10). Nielsen et al. (11)reported the use of a digital filter to reduce noise in cyclic voltammetric mea-
* To whom correspondence should be addressed. 0003-2700/91/0363-2965$02.50/0
surements to detect submillimolar concentrations. Recently, Rice and Nicholson (12) showed that a dopamine concentration of 34 nM can be detected with cyclic voltammetry used in a repetitive mode by the use of ensemble averaging and digital smoothing techniques. Attempts to measure catecholamine neurotransmitters in the brain with voltammetric electrodes require the use of several strategies to improve the quantitative and qualitative aspects of these measurements. Initial efforts were confounded by the large number of substances that were simultaneously detected (13-17). The use of a perfluorinated ionomer (Nafion, a cation-exchange membrane) coated over the electrode tip improved the selectivity and sensitivity for catecholamines over interferant anions such as dihydroxyphenylacetic acid, ascorbate, and uric acid (18). Microelectrodes enabled the use of rapid cyclic voltammetric measurements (19,20),which allowed concentration changes to be monitored in vivo with subsecond time resolution (21). Although the background current is very large at fast scan rates, digital subtraction of the background enables well-defined cyclic voltammograms to be obtained. These can be used to eliminate a large number of molecular candidates from consideration and aid in identification of the detected substance (21,22). With this technique, submicromolar changes in dopamine concentration have been detected in vivo (23, 24). Fast-scan voltammetry has been used in vivo to measure transient changes in catecholamine concentration, such as those which occur during stimulation of neurons (25). Typically, the voltammetric electrode is placed in the terminal field of catecholamine-containing neurons, and the neurons are stimulated with an electrical-pulse train delivered with a separate electrode placed near the axons of the appropriate neurons (26-29). The pulses cause the dopamine-containing neurons to depolarize in synchrony and secrete the neurotransmitter. The amplitude of the current for the oxidation of catecholamines obtained from successive voltammograms provides a measure of the time course of the concentration change during and after the stimulation. For the specific case of detection of dopamine in a brain region known as the caudate nucleus, the concentration change 0 1991 American Chemical Society
